Rapid Raman Imaging of Stable, Functionalized Nanoshells in

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NANO LETTERS

Rapid Raman Imaging of Stable, Functionalized Nanoshells in Mammalian Cell Cultures

2009 Vol. 9, No. 8 2914-2920

Yiming Huang, Vimal P. Swarup, and Sandra Whaley Bishnoi* Illinois Institute of Technology, BCPS Department, 3101 South Dearborn, Chicago, IL Received April 17, 2009; Revised Manuscript Received May 23, 2009

ABSTRACT Two Raman-active poly(ethylene glycol) (PEG) molecules, one linear (MW 5000) and the other branched (MW 2420), are synthesized to stabilize gold-silica nanoshells in cell culture media and track nanoparticles in mammalian cell cultures. The linear PEG provides greater nanoshell stability in saline solution compared to commercially available PEG-thiol or the branched PEG. Surface enhanced Raman scattering rapidly tracks the probes and provides semiquantitative information regarding particle localization within mouse macrophage (RAW 264.7) and human breast cancer (MCF 7) cell cultures.

Metal nanoparticles have found a wide variety of biologically important applications, including use as antimicrobial agents,1,2 biological sensors,3-5 and cancer therapy agents.6-10 Tracking nanoparticle interactions with cells and biological tissues is often done using electron microscopy;11 however, sample preparation can be tedious and the throughput is often low. More recently, surface enhanced Raman scattering (SERS) based imaging techniques have shown promise for their ability to image a large number of cells in a relatively short amount of time.12-14 Since SERS imaging does not require long measurement times and gives narrow and intense vibrational peaks, it has demonstrated the potential for rapid multiplexed imaging.9,12 Some of the benefits of SERS imaging are the ability to monitor particle uptake in situ without significant sample preparation, the quantitative nature of SERS, and the ability to create multifunctional SERS probes. The SERS probes used for this technique are photostable and the wavelengths chosen for Raman excitation are often in the near-infrared spectral region where the background contribution from biological tissues is minimized. Therefore, SERS imaging provides highly specific and quantitative information that allows rapid screening of nanoparticle uptake in cell cultures.15 Multifunctional probes often consist of a Raman-active dye, a targeting moiety, and a stabilizing polymer such as poly(ethylene glycol) (PEG). The stability of nanoparticles in media and saline environments is an important factor in determining their biological behaviors in vivo and in vitro. Different strategies have been employed to provide longterm solution stability of SERS probes in vivo, including * To whom correspondence should be addressed. E-mail: [email protected]. 10.1021/nl901234x CCC: $40.75 Published on Web 07/02/2009

 2009 American Chemical Society

the use of core-shell systems. One such system, popularized by Oxonica Inc., consists of a gold nanoparticle upon which a self-assembled monolayer of Raman-active dyes is attached.12 These particles are then encapsulated within a silica matrix that can be subsequently modified with targeting species and PEG. A second core-shell geometry uses Raman-active dyes embedded within silica matrices upon which metal nanoparticles are then attached.14 Other groups have relied upon electrostatic interactions between the SERSactive dye and the particle surface.15-17 To date, two successful applications of multifunctional Raman-active dyes are the 5,5′-dithiobis(succinimidyl-2-nitrobenzoate) (DSNB) linking groups utilized by Porter, et al.18 for the attachment of antibodies to gold metal surfaces and the recent incorporation of Raman-active dyes into short PEG chains by Graham, et al.19 However, the ability of these PEG chains to stabilize particles in saline has not been compared to commercially available PEG-thiols. One of the challenges in using PEG coatings for nanoparticle stabilization is providing a quantitative analysis of the number of ligands attached to each particle. Previously, Raman active PEG monolayers have been created to estimate the packing density of PEG on a nanoshell surface. These are synthesized through the reaction of p-mercaptoaniline (pMA) with commercially available n-hydroxylsuccinimide labeled PEG molecules.20 However, the ability of these ligands to provide saline stability has not been evaluated to date. In this work, we investigate the stabilizing capabilities of two different Raman active PEG geometries, a linear pMA-PEG (MW 5000) (Scheme 1a) and a branched pMAPEG3 (MW 2420) (Scheme 1b). Nanoshells coated with these Raman-active PEG layers are also used for semiquantitative

Scheme 1. Synthesis of (a) Linear Raman-Active PEG (pMA-PEG) and (b) Branched Raman-Active PEG (pMA-PEG3)

tracking of nanoshells with two different mammalian cell types, human breast cancer (MCF 7), and mouse alveolar macrophage cells (RAW 264.7). Here, we determine that linear Raman-active PEGs enhance particle stability in saline solutions compared to either traditionally used PEG-thiol or the branched Raman-active PEGs. Gold-silica nanoshell stabilization and delivery are the focus of this study due to the significant Raman scattering enhancement of nanoshells without the need to preaggregate the particles.21 However, the strategy developed here can easily be extended to tracking the uptake of other plasmonic nanoparticles, such as silver and gold colloids. Nanoshells are synthesized according to published methods.22 The final particle hydrodynamic diameter (Zave) and ζ-potential are 197 nm and -31 ( 1 mV, respectively, based on dynamic light scattering and ζ-potential measurements (Zetasizer Nano, Malvern, UK). The wavelength of maximum absorption (V-530, Jasco, Japan) of the gold-silica nanoshells used in these studies is 710 nm with significant absorption at 785 nm (the wavelength of the pump laser used in the Raman studies), increasing the coupling between the electromagnetic field of the laser with the particle’s electromagnetic field. To synthesize the linear Raman active PEG (pMA-PEG), methoxypolyethylene glycol succinate N-hydroxysuccinimide (NHS-PEG(5000)) (5000MW, Sigma, 65 mg, 0.013 mmol) is dissolved in 1 mL of 1 mM phosphate buffer (pH 9.1) and added slowly to p-mercaptoaniline (pMA, 125.19 MW, Sigma, 5 mg, 0.04 mmol) in 50 µL of dimethyl sulfoxide (DMSO) and reacted overnight. The resulting product is placed into a 2K MWCO dialysis cartridge (Slide-A-Lyzer, Pierce) and dialyzed against 100 mM, pH 8.0 phosphate buffer to remove excess pMA. The same strategy is used to synthesize the branched Raman-active PEG (pMA-PEG3) except replacing the NHS-PEG (5000) with (methyl-PEG12)3PEG4-NHS Ester (TMS-PEG12, Pierce, MW 2420.8). The concentrations of the final products are obtained by measuring the absorption of pMA in buffer using a molar extinction coefficient of 31694 M-1cm-1 at 200 nm. To test modified particle stability, PEG-thiol (5000 MW, Sigma, St. Louis, MO), pMA-PEG (5000), and pMA-PEG3 (2420) are added into 1 mL of nanoshells (Abs710 nm ) 1.16, ∼3 × 109 Nano Lett., Vol. 9, No. 8, 2009

particles/mL) at a final PEG concentration of 2.65 × 10-6, 2.65 × 10-7, and 2.65 × 10-8M. After an overnight incubation period, 100 µL of 9% sodium chloride (NaCl) is added to the solution to test saline-induced flocculation. This concentration of saline was used because it represents the overall ionic strength used for in vivo experiments with PEGylated nanoshells.23 UV-vis spectrophotometry is used to determine the degree of nanoparticle aggregation through the decrease in the plasmon absorption at 710 nm (Figure 1). If the absorbance change of the solution at 710 nm is less than 10% after 2 h in saline, then the particles are considered to be stable. A broadening of the nanoshell peak and a red shift in the wavelength is a sign of insufficient particle stability. The ratio of (A710 after saline)/(A710 before saline) × 100 was used to assess stability (Figure 1a). As expected, nanoshell stability in 0.9% saline increases markedly with increasing PEG input concentration for all PEG configurations tested. The result of triplicate measurements demonstrate that 92 ( 6% of pMA-PEG (5000) coated nanoshells are stabilized at a concentration of 2.65 × 10-6 M, while only 82 ( 2 and 71 ( 10% of individual nanoshells remain in solution after treatment with the same concentration of PEG-thiol (5000) and pMA-PEG3 (2420), respectively (Figure 1A). Decreasing the input concentration of all of the PEGs below 2.65 × 10-7 M results in significant particle aggregation at this ionic strength. It is concluded that pMAPEG (5000) provides the greatest particle stabilization in saline solution. Two effects can account for the lack of stability obtained with the branched pMA-PEG3, (1) reduced surface density due to steric hindrance of the branched polymer and (2) a shorter chain length of the PEG (2420) versus the PEG (5000). Many groups have demonstrated that both of these conditions can lead to a reduction in colloidal particle stability.24 The enhanced stability of pMA-PEG (5000) over PEG-thiol (5000) may be due to π-stacking between the benzene rings of adjacent pMA-PEG (5000) molecules that allows more efficient molecular packing on the nanoparticle surface compared to the typical “brush” configuration of PEG-thiol (5000).20 ζ-potential measurements of each set of polymers with increasing concentration suggest that as PEG molecules bind to the nanoshell surface 2915

Figure 1. (a) Comparison of nanoshell stabilization as a function of PEG composition and PEG concentration after exposure to 0.9% saline for 2 h. (b) ζ-potential measurements of gold-silica nanoshells after 1 h modification with different PEG compositions and concentrations.

Figure 2. (a) Raman spectra of pMA-PEG (5000) at two concentrations (2.6 × 10-7 and 2.6 × 10-6 M) and PEG-thiol at 2.6 × 10-6M. (b) Raman spectra of pMA-PEG3 (2420) at 2.6 × 10-7 and 2.6 × 10-6 M.

they displace negative counterions such as carbonate, reducing the absolute magnitude of the ζ-potential of the particles (Figure 1B). For example, the ζ-potential of pMA-PEG (5000) modified nanoshells changes from -31 to -14.6 mV at the highest concentration used (2.65 × 10-6 M), while the final ζ-potential of the branched pMA-PEG3 (2420) modified nanoshells is only -21.8 mV at the same concentration. These results support our assertion that the coverage from the branched PEG is significantly lower than either of the linear PEGs tested and that pMA-PEG (5000) displaces more negative ions than PEG-thiol (5000) at 2.65 × 10-6 M, leading to its enhanced stability. The enhancement of Raman vibrational modes on metal surfaces is a function of the surface concentration of a molecule and its orientation on the surface.25-27 Previous SERS studies with gold-silica nanoshells have shown that the intensity of pMA on the nanoshell surface is a function of pMA concentration,21 which can be used to quantify the concentration of pMA-PEG layers on nanoshells.20 Raman spectra and images are collected using an inVia microscope (Renishaw, UK) with a 50× objective with excitation from a semiconductor diode laser (785 nm, 270 µW power at the sample). A 10 s acquisition time is used for Raman imaging (static 2916

scans centered at 1050 cm-1), while 30 s acquisition times are used for extended scans (600-1800 cm-1). Though the longer acquisition times provide better signal-to-noise, 10 s collection times provide sufficient S/N for mapping experiments and allow a single cell to be mapped in a more practical time frame. The average acquisition time is 45 min for a cellular image (10 s static scans, 1 µm steps, ∼20 × 20 µm mapping area).19 The Raman spectra of pMA-PEG (5000) at various concentrations (Figure 2a) demonstrate the major vibrational modes of pMA at 1009, 1077, 1178, and 1585 cm-1. In comparison, PEG-thiol has no significant Raman signal, while pMA-PEG3 (2420) (Figure 2b) has the same feature peaks as pMA-PEG (5000). As the pMA-PEG concentration is increased in Figure 2a, little change is seen in the Raman intensity principally because the surface coverage has already beginning to become saturated at 2.6 × 10-7M. This agrees with the ζ-potential measurements for the linear pMA-PEG seen in Figure. 1b in which there were only minor changes in the ζ-potential with a change in the concentration of the linear pMA-PEG at concentrations above 2.6 × 10-7M. However, as the concentration of the branched pMA-PEG3 is increased, a dramatic change in intensity is seen in the SERS intensity Nano Lett., Vol. 9, No. 8, 2009

Figure 3. (a) Bright-field and Raman image (integration between 1070-1090 cm-1) overlay of pMA-PEG3 (2420) coated nanoshells with MCF-7 breast cancer cells after 2 h of incubation. Signal from the pMA-PEG3 (2420) on nanoshells is represented by red false coloring corresponding to the intensity of the SERS signal. (b,c) Control spectra of MCF-7 cells with plain nanoshells and without nanoshells, respectively. Scale bar ) 10 µm.

Figure 4. (a-c) Bright-field images of macrophage cells (a) with plain nanoshells, (b) PEG-thiol (5000) modified nanoshells, and (c) pMA-PEG (5000) modified nanoshells. Images collected at 180° with a 50× objective (NA ) 0.74). Scale bar ) 10 µm. (d-f) Overlay of the bright-field (a-c) and Raman images based on the integration of Raman intensity from 1070-1090 cm-1. (g-i) Representative Raman spectra of samples a-c taken with a 50× objective with an integration time of 10 s (270 µW at sample).

(Figure 2b). This agrees with the dramatic increase in the ζ-potential for this PEG configuration as the concentration is increased from 2.6 × 10-7 to 2.6 × 10-6M (Figure. 1b). Nano Lett., Vol. 9, No. 8, 2009

Based on the strong SERS signals produced from nanoshells modified with pMA-PEG, we expect that these SERS probes can be used to track PEG-modified nanoshells in cell cultures. 2917

Figure 5. Macrophage cells incubated with pMA-PEG (5000) modified nanoshells imaged after incubation times of 1, 2, and 3 h. (a-c) Bright-field images of macrophage cells with nanoshells. (d-f) Overlay of bright-field and Raman images (signal to baseline intensity from 1070-1090 cm-1). The arrows in (d) indicate the presence of weak but perceptible signal from pMA-PEG (5000).

We began our initial testing of nanoparticle tracking using the branched pMA-PEG3 (2420) labeled nanoshells. The MCF 7 cells are maintained in RPMI 1640 media (ATCC, Manassas, VA) with 10% fetal bovine serum, 2 mM L-glutamine and 1% antibiotic solution and kept in a 37 °C humidified atmosphere of 5% CO2. Prior to testing with nanoparticles, cells are plated and grown to 90% confluence on 8-well Nunc Lab-Tek II CC2 Chamber slides, and then incubated for 2 h with pMA-PEG3 (2420) modified particles suspended in RPMI 1640 media. Excess nanoshells are removed, and then the cells are rinsed with phosphate buffered saline to remove weakly adherent nanoshells, fixed with gluteraldehyde, and imaged with the Raman microscope (Figure 3). After a 2 h incubation between the branched pMA-PEG3 (2420) coated nanoshells and MCF 7 cells, the particles are seen to deposit on the outside of the cells from their scattering profiles (orange/pink particles in the brightfield images) (Figure 3a). These samples are then mapped by Raman microscope with 50× objective, 0.24 mW Laser power and 10 s acquisition time for each reading. The intensity map for peak at 1070-1090 cm-1, corresponding to pMA-PEG3 (2420), is overlaid upon the bright-field image of the associated cell. The benefit of using Raman imaging to track the particles is the ability to detect the nanoparticle at the junction between cells, where the scattering from the cell surface prevented clear detection of the particles. The Raman signal is shown to be specific for the SERS probe (Figure 3a) and not present on bare nanoshells (Figure 3b) or the cells themselves (Figure 3c). Since we know that the pMA-PEG3 coating provides less particle stability, we expected to detect at least some signal from the particle system on the cell surface. Testing with the MCF 7 cell line provides 2918

a good example of how these PEGylated nanoparticles will behave in a typical cell culture environment where targeting is not present and nanoparticle uptake is minimal. Mouse macrophage cells (RAW 264.7, ATCC, Manassas, VA) are used to evaluate the capability of the linear pMAPEG (5000) modified nanoshells for in vivo tracking of nanoparticle uptake. Since the role of macrophage cells in vivo is to defend the body against infection through the phagocytosis (uptake) of invading microorganisms, we expect to see nanoshells not only on the periphery of the cell, as seen in MCF-7 cells, but also within the cell body. In addition it was previously demonstrated that macrophage cells have the ability of taking up nanoshells and delivering them into tumor regions.28 The pMA-PEG (5000) modified particles are suspended in DMEM media (ATCC, Manassas, VA) and then incubated with macrophage cell cultures grown on 8-well Nunc Lab-Tek II CC2 Chamber slides (37 °C, 5% CO2) for 2 h. The cells are then rinsed with phosphatebuffered saline to remove weakly adherent nanoshells, fixed with gluteraldehyde, and imaged with the Raman microscope (Figure 4). The morphology of cells incubated with plain nanoshells (Figure 4a) is consistent with that of cells exposed to both PEG-thiol (5000) modified nanoshells (Figure 4b) and pMA-PEG (5000) modified nanoshells (Figure 4c). From the bright-field images, it is obvious that significantly less background binding of nanoshells is present with the PEGmodified nanoshells, as determined by significantly less scattering of the particles as discussed previously. This is expected due to the ability of PEG to stabilize the nanoshells in media and prevent nonspecific precipitation or binding to extracellular matrix proteins. The detection of nanoshells on the cell surface using particle scattering alone can be Nano Lett., Vol. 9, No. 8, 2009

challenging due to the scattering of light from the cells themselves, whereas the detection of the Raman-active PEGmodified nanoshells is trivial using the signal from the pMAPEG (5000) (Figure 4f). Macrophage cells exposed to either plain nanoshells (Figure 4d,g) or PEG-thiol (5000) modified nanoshells (Figure 4e,h) do not exhibit significant background Raman signals and therefore particle interactions with cells cannot be tracked without modification with a Raman-active probe. To further test the efficacy of these probes, nanoshell internalization within macrophages is examined by varying the dosage time interval. RAW 264.7 cells are treated with pMA-PEG functionalized nanoshell solution containing 1.5 × 109 nanoshells mL-1. Forty thousand cells are seeded in different wells of chamber slide. Incubation time with nanoshells is varied in different wells from 15 min to 3 h. During the treatment, the chamber slide is shaken at low speed to prevent nanoshell sedimentation. After the treatment, excess nanoshells are removed by rinsing with PBS and cells fixed with 1% glutaraldehyde. For incubation times less than 1 h, no pMA-PEG signal is detected at the periphery or inside of the macrophage cells. However, at a time point of 1 h, a weak but perceptible signal is found for the pMA-PEG (5000) indicating the presence of nanoshells at the periphery of the macrophage cell (Figure 5d, arrows). In comparison, a significantly greater signal from the SERS probe is detected inside cells treated with nanoshells for the 2 (Figure 5e) and 3 h time intervals (Figure 5f). The increase in Raman signal correlates well with the increase in scattering intensity seen in the bright-field images of the cells at the 2 h (Figure 5b) and 3 h (Figure 5c) time points. Since nanoshells have a large scattering cross section, they are often detectable using scattering based dark-field imaging;29 however, the SERS tracking technique will prove extremely useful with smaller gold and silver nanoparticles that do not have the large scattering cross sections that nanoshells possess. This technique will be easily transferable to other plasmonic particle systems that are being investigated as drug delivery and cancer therapy agents. In conclusion, the linear pMA-PEG (5000) stabilizes nanoshells better than a branched Raman-active PEG (pMAPEG (2420)). In addition, it provides nanoparticle stabilities that are comparable to those found with commercially available PEG-thiols of the same molecular weight. Lower concentrations do not prevent the aggregation of nanoparticles in saline solutions due to an incomplete surface coverage. Both branched and linear pMA-PEGs provide the ability to act as SERS probes since the benzene ring is directly conjugated to the surface of metal. With Raman mapping, we can measure particle uptake within a cell culture in a semiquantitative manner that may prove useful in future nanoparticle toxicity studies. This technique is significantly faster than electron microscopy techniques, allowing one to screen a large number of cells in a relatively short amount of time. The technique can easily be extended to other metal nanoparticle systems that react readily with thiol groups (gold and silver colloids, gold nanorods, etc.). This new method provides a convenient technique for imaging mammalian cell Nano Lett., Vol. 9, No. 8, 2009

cultures containing metal nanoshells stabilized with PEG. Using a mixed monolayer system that includes a targeting agent along with this SERS-probe, this technique can be extended to other medical detection applications and the tracking of cancer nanotherapeutics in vivo. Acknowledgment. The authors thank Professor Rajendra Mehta for access to cell culture facilities and Dr. Genoveva Murrillo for cell culture training and trouble shooting. In addition, the authors thank Scott Kane and Dr. Anastasia Morfesis of Malvern Instruments for the loan of the Zetasizer Nano. Financial support was provided through start-up funds by the College of Science and Letters at IIT. Y.H. and V.P.S. were both supported by Starr Fellowships provided by the Graduate College at IIT. Supporting Information Available: Additional scheme has been added showing the relative size and structures of PEG-thiol (5000), pMA-PEG (5000), and pMA-PEG3 (2420) on the surface of gold-silica nanoshells. The length of the PEGs are drawn approximately to scale. This material is available free of charge via the Internet at http://pubs.acs.org. References (1) Panacek, A.; Kvitek, L.; Prucek, R.; Kolar, M.; Vecerova, R.; Pizurova, N.; Sharma, V. K.; Nevecna, T.; Zboril, R. J. Phys. Chem. B 2006, 110 (33), 16248–16253. (2) Sondi, I.; Salopek-Sondi, B. J. Colloid Interface Sci. 2004, 275 (1), 177–182. (3) Anker, J. N.; Hall, W. P.; Lyandres, O.; Shah, N. C.; Zhao, J.; Duyne, R. P. V. Nat. Mater. 2008, 7, 442–453. (4) Curulli, A.; Valentini, F.; Padeletti, G.; Cusma, A.; Ingo, G. M.; Kaciulis, S.; Caschera, D.; Palleschi, G. Sens. Actuators, B 2005, 111, 526–531. (5) Kneipp, J. Nanosensors Based on SERS for Applications in Living Cells. Surf. Enhanced Raman Scattering 2006, 335–349. (6) Fortina, P.; Kricka, L. J.; Graves, D. J.; Park, J.; Hyslop, T.; Tam, F.; Halas, N. J.; Surrey, S.; Waldman, S. A. Trends Biotechnol. 2007, 25 (4), 145–152. (7) Girdhani, S.; Bhosle, S. M.; Thulsidas, S. A.; Kumar, A.; Mishra, K. P. J. Cancer Res. Ther. 2005, 1 (3), 129–31. (8) Huang, X.; El-Sayed, I.; Qian, W.; El-Sayed, M. J. Am. Chem. Soc. 2006, 128 (6), 2115–2120. (9) Qian, X.; Peng, X. H.; Ansari, D. O.; Yin-Goen, Q.; Chen, G. Z.; Shin, D. M.; Yang, L.; Young, A. N.; Wang, M. D.; Nie, S. Nat. Biotechnol. 2008, 26 (1), 83–90. (10) Yu, C.; Nakshatri, H.; Irudayaraj, J. Nano Lett. 2007, 7 (8), 2300– 2306. (11) Hauck, T. S.; Ghazani, A. A.; Chan, W. C. W. Small 2008, 4 (1), 153–159. (12) Keren, S.; Zavaleta, C.; Cheng, Z.; de la Zerda, A.; Gheysens, O.; Gambhir, S. S. Proc. Natl. Acad. Sci. U.S.A. 2008, 105 (15), 5844– 5849. (13) Yu, K. N.; Lee, S. M.; Han, J. Y.; Park, H.; Woo, M. A.; Noh, M. S.; Hwang, S. K.; Kwon, J. T.; Jin, H.; Kim, Y. K.; Hergenrother, P. J.; Jeong, D. H.; Lee, Y. S.; Cho, M. H. Bioconjugate Chem. 2007, 18 (4), 1155–1162. (14) Woo, M.-A.; Lee, S.-M.; Kim, G.; Baek, J.; Noh, M. S.; Kim, J. E.; Park, S. J.; Minai-Tehrani, A.; Park, S.-C.; Seo, Y. T.; Kim, Y.-K.; Lee, Y.-S.; Jeong, D. H.; Cho, M.-H. Anal. Chem. 2009, 81 (3), 1008– 1015. (15) Kneipp, J.; Kneipp, H.; Rajadurai, A.; Redmond, R. W.; Kneipp, K. J. Raman Spectrosc. 2009, 40, 1–5. (16) Sajja, H. K.; East, M. P.; Mao, H.; Wang, Y. A.; Nie, S.; Yang, L. Curr. Drug DiscoVery Technol. 2009, 6 (1), 43–51. (17) Gao, X.; Cui, Y.; Levenson, R. M.; Chung, L. W.; Nie, S. Nat. Biotechnol. 2004, 22 (8), 969–76. (18) Grubisha, D. S.; Lipert, R. J.; Park, H. Y.; Driskell, J.; Porter, M. D. Anal. Chem. 2003, 75 (21), 5936–5943. (19) McKenzie, F.; Ingram, A.; Stokes, R.; Graham, D. Analyst 2009, 134 (3), 549–56. 2919

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Nano Lett., Vol. 9, No. 8, 2009